Comparative Biochemistry and Physiology Part A 120 (1998) 469–475

In vivo 31P-NMR spectroscopy and respiration measurements ofanaesthetized goby (Pomatoschistus sp.) pre-exposed to ammonia

Jon Arne Grøttum a,*, Ulf Erikson b, Hans Grasdalen b, Magne Staurnes a

a Department of Zoology, Norwegian Uni6ersity of Science and Technology, N-7055 Drag6oll, Norwayb Department of Biotechnology, Norwegian Uni6ersity of Science and Technology, N-7034 Trondheim, Norway

Received 12 November 1996; received in revised form 8 March 1998; accepted 24 March 1998

Abstract

The effects of ammonia pre-exposure on high energy-phosphates and the intermediary metabolism of the marine teleost Goby(Pomatoschistus sp.) were studied by in vivo 31P-NMR with simultaneous respirometry. No significant effect of ammoniapre-exposure was observed on oxygen uptake, muscle intracellular pH or high-energy phosphate metabolism. Brief metomidateanaesthesia (10 mg l−1) followed by continuous light anaesthesia (1 mg l−1) was used to prevent excessive swimming. This hadno effect on oxygen uptake, muscle intracellular pH or high-energy phosphate metabolism. © 1998 Elsevier Science Inc. All rightsreserved.

Keywords: Ammonia; Toxicity; Marine fish; High energy-phosphate; Intracellular pH; Oxygen consumption; 31P-NMR; Anaesthe-sia

1. Introduction

The specific biochemical mechansim of ammonia tox-icity is associated with effects on carbohydrate and/or amino acid metabolism [1,5,11,16,27]. Via its effectson intermediary metabolism, acute ammonia expo-sure results in a depletion of energy stores (includingphosphocreatine (PCr) and ATP) in the brainstem[1,4,28].

Changes in intermediary metabolism of ammonia-ex-posed fish include a decrease in glutamic acid dehydro-genase (GlDH) activity and ATP levels in brain,erythrocytes, gill, liver and plasma [1,4,16,19,29]. This isexplained by an increase in glutamine synthetase activ-ity, which plays an important role in the detoxificationof free ammonia. The following reaction is suggested byBuckley et al. [6] and Mehre et al. [23].a-keto-glutaric acid+NADH+NH4

+ lGIDHglutamic acid

+NAD+H2O

Glutamic acid+ATP+NH4+ lGlutamine synthetase

glutamine+ADP+Pi

The detoxification reaction may impair the energybalance in the organism due to ATP consumption byglutamine synthesis. Increased GlDH activity also with-draws a-ketoglutaric acid from the tricarboxylic acidcycle [13], decreasing the capacity of the cycle. TheNADH level falls along with the increase in GlDHactivity which in turn affects mitochondrial electrontransport.

It has been speculated that since ammonium ionsaffect several key enzymes in energy metabolism, ele-vated ammonia levels might reduce the capacity ofmuscle to exercise or affect the nervous co-ordinationof exercise, either centrally or by disrupting peripheralmotor innervation [3]. White muscle ATP and totaladenylate concentrations as well as adenylate energycharge have been regarded as useful indicators of envi-ronmental stress in several fish species [40]. MacFarlane[20] reported lower values of ATP and adenylate energycharge (AEC=(ATP+1

2ADP)(ATP+ADP+AMP)−1)* Corresponding author. Tel.:+47 73597785; fax: +47 73596311;

e-mail: jon.arne.grottum@chembio.ntnu.no

1095-6433/98/$19.00 © 1998 Elsevier Science Inc. All rights reserved.PII S1095-6433(98)10047-8

J.A. Grøttum et al. / Comparati6e Biochemistry and Physiology, Part A 120 (1998) 469–475470

in brain, gill and liver and most notably, in muscletissues when gulf killifish (Fundulus grandis) were ex-posed to sublethal pH levels for 96 h.

On the basis of our knowledge of the toxic effect ofammonia on intermediary metabolism, we studied thepossibility of using muscle high-energy phosphates(HEP) and intracellular pH (pHi) as indicators of sub-lethal effects of ammonia on fish. As biochemicalchanges precede cellular and systematic changes, theymay be detected much earlier than alterations in growthchanges, deformity, diseases, mortality etc. In vivo 31P-NMR spectroscopy in combination with simultaneousrespiration measurements, therefore, seemed to be anattractive method for studying the possible effects ofammonia exposure on fish. In vivo NMR has severaladvantages as it is non-invasive and non-destructive,while tissue-sampling and extraction artefacts are elimi-nated [37]. In vivo 31P-NMR has been used to studyenergy metabolism in several freshwater fish species,including effects on the energy metabolism of loach(Cobitis biswae) [9,10], on the acid–base metabolism ofacid-exposed tilapia (Oreochromis mossambicus) [39], onacidosis in anoxic goldfish (Carassius auratus), tilapiaand common carp (Cyprinus carpio) [38] and duringhypoxia and recovery of the latter three species [37].

In the case of marine species, the method has beenused to study the sublethal effects of different environ-mental pollutants on blue mussels (Mytilus edulis).Several pollutants were found to give distinct reductionsin high-energy phosphates [2]. In red abalone (Haliotisrufescens) hypoxia induced a decrease in phospho-arginine and pHi while ATP remained unaffected dur-ing 1 h tidal exposure to air [33].

Goby (Pomatoschistus sp.) was used because of itsrelatively cylindrical body shape, which makes it suit-able for use in NMR tubes and because of its availabil-ity. The fish live in shallow seawater and can be foundat night in tidal pools that are often exposed to subop-timal environmental conditions [15,26].

The primary aim of this study was to examinewhether ammonia exposure affected intermediarymetabolism, observed as changes in muscle phospho-metabolites and pHi. Our second objective was to eval-uate whether 31P-NMR might be a usefulmethod to monitor any effects on muscle metabolism ofenvironmental stressors such as adverse water quality.

2. Materials and methods

2.1. Experimental protocol

Goby (Pomatoschistus sp.) weighing 0.3–0.9 g werecaught in the Trondheimsfjord in Central Norway. Thefish were kept in an aquarium with a continuous supplyof fresh filtered seawater (34% salinity, 1091°C) for 14

days prior to the experiment.A preliminary experiment was carried out to deter-

mine suitable levels for subacute ammonia exposure.Five groups of three fish in 1 l seawater were exposed to2, 4, 6, 8 and 10 mM total ammonia [NH3+NH4

+](TA) with pH 7.8–7.9. After 24 h, 100% mortality wasobserved in the groups exposed to 6, 8 and 10 mM TA,while no mortality occurred in the groups exposed to 2and 4 mM. The highest level without mortality (4 mM)was used for a 24 h exposure.

The ammonia-exposed group and the control groupwere kept in plastic tanks (15 l, 1091°C) with aeratedseawater. When studying the effects of ammonia expo-sure, we were left with a dilemma as to whether anyeffects on metabolism might be due to TA exposure perse, or might originate from struggling due to escapebehaviour after recovery from initial anaesthesia. Tominimise the latter possibility, we chose to use lightanaesthesia during the entire experimental period.Therefore, each fish was anaesthetized with metomidate(1 - (1 - phenylethyl) - 1H - imidazole - 5 - carboxylic acidmethyl ester hydrochloride) (Marinil™, Wildlife Labs,Fort Collins, CO) (10 mg l−1) in seawater (controlgroup, n=4) or in seawater with TA (4 mM) (pre-ex-posed group, n=4) before transfer to the spectrometer.When each fish was quiet and did not respond on touch(after 2.5–4.0 min), it was transferred to the NMR tube(10 mm diameter) (flow cell) containing metomidate (1mg l−1) in seawater without (control group) or with TA(4 mM) (pre-exposed group). Seawater without (controlgroup) or with TA (4 mM) (pre-exposed group) circula-tion through the flow cell was started and the cell wasplaced in the spectrometer magnet 5–8 min after startof anaesthesia. The flow cell is shown in Fig. 1. Thebottom of the cell was packed with cotton (10 mm) inorder to ensure that the thickest (central) part of the fishwas located in the rf coil in the spectrometer magnet.The flow cell was supplied with the same seawater asmentioned above from reservoirs (250 ml flasks withaeration, 100% O2 saturation) placed in a water bath(10°C). The water was pumped (Gilson Minipuls 3)(1.27 ml min−1) through a flexible tube (Masterflex®

Tygon® tubing, size 13 (Gas permeability O2; 0.7–12(cm3×mm)× (s×cm−2×cmHg)−1×10−10). A simi-lar tube was used from the flow cell to the oxygenelectrode. To ensure constant water level (55 mmheight), the flow cell was fitted with a slurp tube. Thelength of the tubes from the magnet to the reservoir was1.8 m. After the final spectrum had been taken it wasverified from opercular movements that the fish werestill alive.

2.2. In 6i6o 31P-NMR measurements

Each spectrum was accumulated at �8°C over a

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Fig. 1. Flow-cell assembly. Experimental setup used for in vivo 31P-NMR combined with respiration measurements. A, spectrometer magnet; B,flow-cell; C, fish; D, water-supply tube; E, slurp tube; F, outlet tube; G, oxygen electrode; H, oxygen meter; I, peristaltic pump; J, seawaterreservoir; K, aerator; F, water bath.

period of 5–19 min from a cross section of the centralportion of the whole fish, i.e. NMR-signals originatedpredominantly from white muscle. A JEOL EX-400spectrometer operating at 161.7 MHz for 31P was used.Sample tubes were not spun and no field/frequency lockwas applied. A spectral width of 8 KHz, 33 K datapoints, a 60° pulse angle, a pulse repetition time of 4 sand nuclear overhauser enhancement with suppressionand inverse-gated proton decoupling were used. PCrwith a resonance at −3 ppm in living tissue served asa standard for the chemical shift measurements.

The pHi was calculated from the equation givenby Van Ginneken et al. [37] based on the chemical shiftof inorganic phosphate (Pi) relative to PCr reson-ances.

2.3. Respiration measurements

The rate of metabolism was measured by registeringthe oxygen level of the flow-cell water outlet with aRadiometer (E5046) oxygen electrode (Fig. 1). Theelectrode was installed in a glass water jacket withcirculating water (25°C). Oxygen consumption was cal-culated according to Cech [8] and Colt [12], respec-tively. Because of differences in weight, the data werecorrected according to Petersen and Petersen [26](VO2

=a ·bodyweight0.75) to eliminate the allometric ef-fect on metabolism.

2.4. Statistics

A two-tailed Student’s t-test was used [42] to testsignificant differences (P50.05) between the groups inphosphometabolites, pHi and oxygen consumptions.The percentage values of phosphometabolites derivedfrom NMR spectra were arc-sinus transformed beforestatistical analysis. The calculations were performed byusing Systat 5.04.

3. Results

We observed no differences in behaviour, generalappearance or mortality (no mortality) between thecontrol group and ammonia- pre-exposed groups before,during or after the fish were placed in the flow-cell.

Typical 31P-NMR spectra of body cross-sections arepresented in Fig. 2, which show a control fish 20 min.(Fig. 2A) and 40 min (Fig. 2B) after start of anaesthesia.The relative resonance intensity of Pi decreased with timeduring anaesthesia and transfer to the spectrometer. Thiswas also the case for ammonia-preexposed fish.

A semi-quantitative comparison of treatments, inwhich the metabolites are expressed as a percentage oftotal NMR-visible phosphorus, is shown in Table 1. ThePi and sugar phosphates (SP including AMP and

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Fig. 2. In vivo 31P-NMR spectra of goby (Pomatoschistus sp.). A, control fish 20 min after start of anaesthesia; B, control fish 40 min start ofanaesthesia; C, exposed fish 40 min start of anaesthesia.

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Table 1Metabolite levels, intracellular pH and oxygen consumption of the control group and ammonia-preexposed group of gobies (Pomatoschistus sp)

PCr ATP Pi SP Pi: PcrTime PCr:ATP pH VO2

66.093.2 9.690.5 14.891.8 9.693.0Control 0.290.023911 7.090.5 7.3390.04 64.298.949.599.7 10.790.6Pre-exposed 27.698.11794 12.291.1 1.190.7 4.991.1 7.2190.03 46.298.4

0.20 0.30 0.24 0.51 0.31 0.19 0.08 0.30P

All values are mean9SE (n=4). Time indicates the start of the measurement (min after start of anaesthesia). Metabolite levels are given as apercentage of total NMR-visible phosphorus. Oxygen consumption is in mg kg−1 h−1.P is the significance level.

IMP) values of the ammonia- pre-exposed fish seemedhigher with corresponding lower PCr and ATP values.The ammonia- pre-exposed fish thus had an apparentrelatively lower [Pi:PCr] ratio and a lower [PCr:ATP]ratio. The mean pHi values of the control group andammonia pre-exposed group were 7.3 and 7.2, respec-tively. However, none of the differences weresignificant.

The weight-corrected rates of oxygen consumption ofthe groups (Table 1) were not significantly different.

4. Discussion

The spectra of metomidate-anaesthetized fish exhib-ited typical features of unstressed fish, i.e. a small orbarely visible Pi peak and a concomitantly high PCr-peak and a pHi in the physiological range of unstressedfish [10,34–36]. However, it was apparent that induc-tion of anaesthesia or handling associated with transfer-ring the fish to the flow cell had a minor effect onhigh-energy phosphate metabolism since we observed aslight recovery effect over a 20–40 min period duringperfusion in the spectrometer. Metomidate thus did notseem to affect white muscle aerobic HEP which is inaccordance with results reported by Chiba et al. [9]during ethyl-p-aminobenzonate and 2-phenoxyethanolanaesthesia in loach muscle.

Malmstrøm et al. [21] reported that metomidate (]10 mg l−1) anaesthesia caused a more complete relax-ation of Atlantic halibut (Hippoglossus hippoglossus L.)muscle than did MS-222. Anaesthesia of Atlanticsalmon (Salmo salar) using concentrations higher than3 mg l−1 metomidate prevented plasma cortisol re-sponse to handling stress presumably due to a blockageat the interrenal cell level, whereas blood lactate levelsand haematocrit increased [25]. This was confirmed byThomas et al. [32] who reported that metomidate (7 mgl−1) appeared to exert a direct action at the interrenalgland to block ACTH stimulation of steroidogenesis injuvenile red drum (Sciaenops ocellatus). Mattson andRiple [22] considered metomidate (effective concentra-tion 5 mg l−1) to be an efficient anaesthetic for use oncod (Gadus morhua), causing no mortality. Further-more, metomidate-induced sleep rather than general

anaesthesia was reflected in the maintenance of opercu-lar respiration, whereas benzocaine and MS-222 causedreduced ventilation, resulting in hypoxia.

However, it has been reported that prolonged expo-sure to anaesthesia impairs oxygen delivery to tissuesand produces carbon dioxide accumulation, due toincreased erythrocyte membrane permeability, whichchanges the plasma [Na+: K+] ratio and causes cellswelling and increased blood haematocrit. Further-more, the blood pH falls because of lactic acidosis [14].Since we did not observe serious acidosis (pHi 7.2–7.3)and since it has been reported that metomidate doesnot affect respiration [22], we assumed the fish were ina truly physiologically relaxed state during the experi-mental period of a maximum of 40 min. It may bespeculated whether the anaesthetic might have had aquick recovery effect on the HEP metabolism of theammonia-preexposed fish, i.e. before the first spectrumwas acquired. However, recovery from severe perturba-tions of metabolism was not very likely since typicalrecovery times of PCr, ATP and pHi from anoxia are inthe range of 3–4 h [35,39] or even more after exhaust-ing exercise [17]. Taken together, these studies essen-tially complement our results, that show that briefmetomidate anaesthesia followed by continuous lightanaesthesia probably do not affect oxygen uptake, mus-cle pHi or HEP.

To our knowledge, no information is available re-garding HEP in white muscles during ammonia expo-sure. The mean HEP values of our ammonia-exposedfish were apparently lower than of those of the controlfish. The differences, however, were not significant.Ammonia toxicity has been reported as affecting inter-mediary metabolism in other parts of the body such asthe gills, liver, erythrocytes and plasma [4–6,16,18,24,27,31]. In muscle tissues, the ATP and theadenylate energy charge of acid-exposed gulf killifishhave been reported to decrease significantly [20]. How-ever, Van Waarde [39] reported no significant depletionof tissue PCr and ATP when tilapia were exposed tolow water pH.

The mean pHi of the ammonia-exposed fish wasapparently lower, but not significantly, than that of thecontrol fish. To our knowledge no observations havereported effects on pHi as a result of ammonia expo-

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sure. However, ammonia is reported to increase intra-cellular levels of lactate in blood, brain and gill[1,4,19,30], which indicates anaerobic metabolism andpossible acidification of the tissue. In contrast to theseobservations, non-respiratory alkalosis was observed ina rainbow trout (Oncorhyncus mykiss) in seawater [41].In fully relaxed fish, i.e. when the Pi peak was barelyvisible, the pHi was between 7.3 and 7.4, which indi-cates that our fish were close to complete relaxation[35,37].

Nemcsok et al. [24] reported that NH3 exposureinduced anoxia in carp (Cyprinus carpio), silver carp(Hypophthamicthys molitrix) and sheatfish (Silurusglanis). Furthermore, [35,37,38] reported decreasedoxygen consumption, reduced PCr and pHi levels andincreased Pi levels for anoxic carp (Cyprinus carpio),tilapia (Sarotherodon mossambicus) and goldfish(Carassius auratus). Since we did not observe theseeffects, no white muscle tissue anoxia apparently oc-curred in our experiment.

No significant differences between the two groupswere observed on the HEP metabolism or pHi, which isin accordance with the insignificant difference observedin oxygen consumption. Oxygen values were compara-ble with the data reported by Petersen and Petersen[26], who found a routine consumption rate of 48–72mg kg−1 h−1 for sand goby (Pomatoscistus minutus)(1–3 g, 6°C).

Our study suggests that 31P-NMR studies of fishwhite muscle HEP metabolism to demonstrate environ-mental stressors have certain limitations. In cases wherethe stressors exert a direct effect upon aerobicmetabolism, such studies might be useful. However,NMR studies using a surface coil on the brain or gillsof larger fish in larger flow-cells [36,37,39] may proveuseful. Chemical analysis of liver nucleotides ratherthan in muscle may also be an alternative [7].

Acknowledgements

This work was carried out as a part of the SINTEFStrategic Technology Programme in Aquaculture andfinancially supported by the Research Council of Nor-way (NTNF-project no. 26877).

References

[1] Arillo A, Margiocco C, Melodia F, Mensi P, Schenone G.Biochemical aspects of water quality criteria: the case of ammo-nia pollution. Environ Tech Lett 1981;2:285–92.

[2] Aunaas T, Einarson S, Southon TE, Zachariassen KE. Theeffects of organic and inorganic pollutants on intracellular phos-phate compounds in blue mussels (Mytilus edulis). CompBiochem Physiol 1991;100C:89–93.

[3] Beaumont MW, Butler PJ, Taylor EW. Plasma ammonia con-centration in brown trout in soft acidic water and its relationshipto decreased swimming performance. J Exp Biol 1995;198:2213–20.

[4] Begum SJ. Biochemical adaptive responses in glucosemetabolism of fish (Tilapia mossambica) during ammonia toxic-ity. Curr Sci 1987;56:705–8.

[5] Berl S. Cerebral amino acid metabolism in hepatic coma. ExpBiol Med 1971;4:71–84.

[6] Buckley JA, Whitmore CM, Liming BD. Effects of prolongedexposure to ammonia on the blood and liver glycogen of cohosalmon (Oncorhynchus kisutch). Comp Biochem Physiol1979;63C:297–303.

[7] Caldwell CA, Hinshaw J. Physiological and haematological re-sponses in rainbow trout subjected to supplemental dissolvedoxygen in fish culture. Aquaculture 1994;126:183–93.

[8] Cech JJ. Respirometry. In: Schreck CB, Moyle PB, editors.Methods in Fish Biology. MA, USA: American Fisheries Soci-ety, 1990:1990.

[9] Chiba A, Hamaguchi M, Kosaka M, Asai T, Tokuno T,Chichibu S. In vivo 31P-NMR analysis of the electric anaes-thetised loach, Cobits biswae. Comp Biochem Physiol1990;97A:385–9.

[10] Chiba A, Hamaguchi M, Kosaka M, Tokuno T, Asai T,Chichibu S. Energy metabolism in unrestrained fish with in vivo31P-NMR. Comp Biochem Physiol 1990;96A:253.

[11] Clifford A, Prior R, Hintz H, Brawn P, Visek W. Ammoniaintoxication and intermediary metabolism. Biol Med1972;140:1447–50.

[12] Colt J. Computation of dissolved gas concentration in water asfunction of temperature, salinity and pressure. Am Fish Soc SpecPubl 1984;14:155.

[13] Fahien LA, Hsu SL, Kmiotek E. Effect of aspartate on com-plexes between glutamate dehydrogenase and various amino-transferases. J Biol Chem 1977;252:1250–6.

[14] Heisler N. Acid–base regulation in fishes. In: Heisler N, editor.Acid–Base Regulation in Animals. Amsterdam: Elsevier,1986:309–56.

[15] Jaquet N, Raffaelli D. The ecological importance of the sandgoby Pomatoschistus minutus (Pallas). J Exp Mar Biol Ecol1989;128:147–56.

[16] Jeney G, Nemesok J, Jeney Zs, Olah J. Acute effect of sublethalammonia concentration on common carp (Cyprinus carpio L.).II. Effect of ammonia on blood plasma transaminase (GOT,GPT), GIDH enzyme activity and ATP value. Aquaculture1992;104:149–56.

[17] Kieffer JD, Currie S, Tufts BL. Effects of environmental temper-ature on the metabolic and acid–base responses of rainbow troutto exhaustive exercise. J Exp Biol 1994;194:299–317.

[18] Kuhn B, Jacobasch G, Gerth C, Rapoport SM. Kinetic proper-ties of phosphofructokinase from erythrocytes of rats and rab-bits. I. The influence of potassium and ammonia ions and ofinorganic phosphate. Eur J Biochem 1974;43:437–42.

[19] Korting VW. Untersuchung zur Einwirknung des Ammoniaksauf das Blut der Fische. Wass Abwass Forsch 1969;4:154–9.

[20] MacFarlane RB. Alteration in adenine nucleotide metabolism inthe gulf killifish (Fundulus grandis) induced by low pH water.Comp Biochem Physiol 1981;68B:193–202.

[21] Malmstrøm T, Salte R, Gjøen HM, Linseth A. A practicalevaluation of metomidate and MS-222 as anaesthetics for At-lantic halibut (Hippoglossus hippoglossus L). Aquaculture1993;113:331–8.

[22] Mattson NS, Riple TH. Metomidate, a better anesthetic for cod(Gadus morhua) in comparison with benzocaine, MS-222,chlorbutanol and phenoxyethanol. Aquaculture 1989;83:89–94.

[23] Mehrle PM, Bloomfield RA. Ammonia detoxifying mechanismsof rainbow trout altered by dietary dieldrin. Toxicol Appl Phar-macol 1974;27:355–65.

J.A. Grøttum et al. / Comparati6e Biochemistry and Physiology, Part A 120 (1998) 469–475 475

[24] Nemcsok J, Gyore K, Olah J, Boross L. Effect of NH3 on bloodglucose and catecholamine levels, GOT, GPT, LDH enzymeactivity and respiration of fishes. Symp Biol Hung 1984;23:209–17.

[25] Olsen YA, Einarsdottir IE, Nilssen KJ. Metomidate anaesthesiain Atlantic salmon, Salmo salar, prevents plasma cortisol in-crease during stress. Aquaculture 1995;134:155–68.

[26] Petersen JK, Petersen GI. Tolerance, behaviour and oxygenconsumption in the sand goby, Pomatoschistus minutus (Pallas),exposed to hypoxia. J Fish Biol 1990;37:921–33.

[27] Prior RL, Visek WJ. Effect of urea hydrolysis on tissue metabo-lite concentration in rats. Am J Physiol 1972;223:1143–9.

[28] Schenker S, McCandless DW, Brophy E, Lewis MS. Studies onthe intracerebral toxicity of ammonia. J Clin Invest1967;46:838–48.

[29] Schenone G, Arillo A, Margiocco C, Melodia F, Mensi P.Biochemical bases for environmental adaption in goldfish(Carassius auratus L.): resistance to ammonia. Ectox Env Saf1982;6:479–88.

[30] Shaffi SA. Ammonia toxicity: metabolic disorder in nine fresh-water teleosts. Toxicol Lett 1980;6:349–56.

[31] Sousa RJ, Meade TL. The influence of ammonia on the oxygendelivery system of coho salmon haemoglobin. Comp BiochemPhysiol 1977;58A:23–8.

[32] Thomas P, Robertson L. Plasma cortisol and glucose stressresponses of red drum (Sciaenops ocellatus) to handling andshallow water stressors and anesthesia with MS-222, quinaldineand metomidate. Aquaculture 1991;96:69–86.

[33] Tjeerdema RS, Kauten RJ, Crosby DG. Sublethal effects ofhypoxia in the abalone (Haliotis rufescens) as measured by invivo 31P NMR spectroscopy. Comp Biochem Physiol1991;100B:653–9.

[34] Van Den Thillart G, van Waarde A. The role of metabolic

acidosis in the buffering of ATP by phosphagen stores in fish: anin vivo NMR study. In: Hochachka PW, editor. SurvivingHypoxia: Mechanisms of Control and Adaption. Boca Raton,Lousiana: Chemical Rubber Company, 1993:1993.

[35] Van den Thillart G, van Waarde A, Muller HJ, Erkelens C,Addink A, Lugtenburg J. Fish muscle energy metabolism mea-sured by in vivo 31P-NMR during anoxia and recovery. Am JPhysiol 1989;256:R922–9.

[36] Van den Thillart G, van Waarde A, Erkelens C, Lugtenburg J. Aflow-through probe for in vivo 31P NMR spectroscopy ofunanesthetized aquatic vertebrates at 9.4. Tesla J Magn Reson1989;84:573–9.

[37] Van Ginneken V, van den Thillart G, Addink A, Erkelens C.Fish muscle energy metabolism measured during hypoxia andrecovery: an in vivo 31P-NMR study. Am J Physiol1995;268:R1178–87.

[38] Van Waarde A, de Graaff I, van den Thillart G, Erkelens C.Acidosis (measured by nuclear magnetic resonance) and ethanolproduction in anoxic goldfish acclimated to 5 and 20°C. J ExpBiol 1991;159:387–405.

[39] Van Waarde A, van den Thillart G, Erkelens C. Functionalcoupling of glycolysis and phosphocreatine utilization in anoxicfish muscle. An in vivo 31P NMR study. J Biol Chem1990;265:914.

[40] Vetter RD, Hodson RE. Use of adenylate concentrations andadenylate energy charge as indicators of hypoxic stress in estuar-ine fish. Can J Fish Aquat Sci 1982;39:535–41.

[41] Wilson RW, Taylor EW. Transbranchial ammonia gradients andacid–base responses to high external ammonia concentration inrainbow trout (Oncorhynchus mykiss) acclimated to differentsalinities. J Exp Biol 1992;166:95–112.

[42] Zar JH. Biostatistical Analysis. NJ, USA: Prentice Hall,1996:1996.

.